Arginine vasopressin (AVP), also known as antidiuretic hormone, induces urinary concentration by renal salt and water reabsorption via activation of the vasopressin V2 receptor (V2R) in the thick ascending limb, distal convoluted tubule, and principal cells (PCs) of the connecting tubule (CNT) and collecting duct (CD).1–5 Clinically, V2R antagonists have been recognized as a promising pharmacologic tool for retardation of chronic kidney disorders, such as polycystic kidney disease and diabetic nephropathy.6–12
Antagonists of another vasopressin receptor, the V1a vasopressin receptor (V1aR), are also emerging in the treatment of patients with CKD,13–15 although renal V1aR distribution and function have been studied less in comparison with V2R.12,16,17 Previous studies suggested V1aR localization in renal vessels, the macula densa (MD), and intercalated cells (ICs) of CNT/CD, but a clear segmental and cellular distribution could not be defined owing to poor specificity of available antibodies.16,18,19 Among many other functions, V1aR signaling has been implicated in renal acid-base handling and renin release.18–20 V1aR-deficient mice exhibit hypotension due to suppression of the renin-angiotensin-aldosterone system (RAAS) and show distal renal tubular acidosis (dRTA), probably due to impaired functioning of ICs.18–20 Renal ICs include the proton-secreting type A intercalated cells (A-ICs) and the bicarbonate-secreting type B intercalated cells (B-ICs) as well as an intermediate form of non-A, non-B ICs.21 Modulation of their activity by AVP has been demonstrated in several models, yet the mediating pathways remained debatable.19,20,22–24 Elucidation of direct V1aR-mediated effects in ICs has been challenging due to parallel AVP-induced stimulation of RAAS,18,25 complex paracrine interactions between ICs and PCs,26 and uncertainty regarding basolateral versus luminal AVP action.27
In view of increasing availability of highly selective V1aR antagonists and growing interest in their therapeutic potential in the treatment of chronic kidney disorders, it is necessary to substantiate current knowledge on renal V1aR distribution and function. The aim of our study is to provide detailed information on segmental and cellular V1aR distribution in rodent and human kidneys, with a particular focus on its role in renal acid-base handling. Using in vivo and ex vivo models, we address the hypothesis that V1aR activation contributes to urinary acidification via A-ICs.
An extended methods description is provided in Supplemental Material.
All animal experiments were approved by the German or Danish Animal Welfare Regulation Authority and performed in accordance with the European Union Directive 2010/63/EU on the protection of animals used for scientific purposes. For localization studies, adult (8–12 weeks old) male C57BL/6J mice, V1aR-deficient mice, Wistar rats, and AVP-deficient Brattleboro rats with central diabetes insipidus (DI) were euthanized by in vivo perfusion with 3% paraformaldehyde (PFA) in PBS, and kidneys were processed for immunohistochemistry or immunofluorescence (n=6 animals in each group). Physiologic studies with DI rats (n=16) were performed in metabolic cages with water and chow ad libitum. After 2 hours of adaptation, DI rats were treated with the V1aR agonist (A0–4-67)28,29 or vehicle (0.9% NaCl; n=8 each group) for 4 hours, and urine samples were collected hourly under mineral oil. After recovery for 3 days in normal cages, the treatment groups were inverted; the vehicle-treated animals received the agonist, whereas the agonist-treated rats received vehicle using the same experimental protocol. Effects of three agonist doses (200 ng/kg, 2 μg/kg, and 10 μg/kg body weight intraperitoneally) were tested in this way, with recovery periods of at least 3 days between the experimental settings. Finally, DI rats were treated with vehicle (n=6) or the V1aR agonist (2 μg/kg body weight; n=7) for 2 hours and decapitated under isoflurane anesthesia to collect blood samples. The pH and HCO3− in plasma and urine were measured using an ABL 800 Flex analyzer. Net acid excretion (NAE) was determined by titration with the same experimental setting as described previously.30 Deviating from the original protocol, the method was validated for a reduced urine volume of 500 μl per sample (Supplemental Tables 1–4).
Male adult C57BL/6J (n=13) were anesthetized using a mix of ketamine (10 mg/ml) and xylazine (1 mg/ml). The urinary bladder was catheterized, and a micro-pH electrode was placed in the outflow of the catheter to measure urine pH every 5 seconds. After establishing baseline pH values for 30 minutes, mice received the V1aR agonist (AO-4–67; 2 μg/kg body weight intraperitoneally; n=6) or vehicle (0.9% saline intraperitoneally; n=7), and pH was measured for another 60 minutes. The effects of a V1aR antagonist (CL-14–10231,32; 2 mg/kg body weight intraperitoneally; n=3) versus vehicle (0.9% saline; n=3) were assessed in bladder-catheterized male adult C57BL/6J mice fed with a metabolic acidosis–inducing diet (0.28 M NH4Cl in chow plus 0.5% sucrose in water; n=10) for 3 days before the evaluation.33 Morphologic evaluation of V1aR distribution was studied in a parallel cohort of mice receiving regular (n=4) versus the acidosis-inducing diet for 3 days (n=4); animals were euthanized by in vivo perfusion.
Generation of the Anti-V1aR Antibody
The peptide sequence NH2-CKDSPKSSKSIRFIPVST-COOH from the C-terminal mouse V1aR portion was chosen to generate the anti-V1aR antibody due to its negligibly low homology with the vasopressin V2 and V1b receptors and high conservation between the mouse, rat, and human species. Peptide synthesis, immunization of rabbits, and affinity purification of anti-V1aR antibodies were performed by Pineda Antibody-Service (Berlin, Germany). Specificity tests were performed using kidneys from V1aR-deficient mice or human embryonic kidney (HEK293) cells transfected with V1aR or V1bR.
HEK293 cells were cultured in DMEM medium on coverslips, transfected with GFP-tagged V1aR or control GFP-containing plasmids (pEFGP-N1) using JetPEI transfection reagent, fixed with 3% PFA/PBS for 10 minutes, and processed for double labeling of V1aR and GFP using immunofluorescence. Alternatively, cells were grown in petri dishes and transfected with V1aR, FLAG-tagged vasopressin V1b receptor (V1bR), or control pcDNA3.1 plasmid.34 Cell lysates were precipitated using protein G Sepharose gel (GE Healthcare Life Sciences) and eluates processed for immunoblotting using anti-V1aR or anti-FLAG antibodies (Sigma-Aldrich). Primary inner medullary collecting duct (IMCD) cells were obtained from adult male Wistar rat kidneys using dissection and chemical digestion of renal inner medulla in identical fashion as described previously.35 Cells were grown on permeable filter support systems to full confluence, treated with the V1aR agonist (1.3 μM) or vehicle from the basolateral side for 4 hours, fixed with 4% PFA/PBS, and processed for immunofluorescence.
mRNA was extracted from microdissected nephron segments using an RNA extraction kit (Stratec Biomedical), and cDNA was synthesized by reverse transcription (Tetro Reverse Transcription; Promega). V1aR-specific forward (5′-CAA TGT CCG AGG GAA GAC AG-3′) and reverse primers (5′-GTT GGG CTT CGG TTG TTA GA-3′) were designed, and RT-PCR was performed in an automated thermal cycler (PerkinElmer) using Taq polymerase (GIBCO).
Immunofluorescence, Immunohistochemistry, and Quantitative Analyses
Paraffin-embedded kidney sections (4 μm) were dewaxed, boiled in citrate buffer (pH 6, 6 minutes), washed in TBS, and blocked with 5% skim milk in TBS for 30 minutes. Cultured cells on coverslips were permeabilized for 10 minutes using 0.5% Triton X-100/TBS and blocked with 5% BSA in TBS for 30 minutes. Primary antibodies to V1aR (own antibody), aquaporin 2 (AQP2; sc-9882; Santa Cruz Biotechnology, Dallas, TX), pendrin (a gift from C.A. Wagner, Zurich, Switzerland), vacuolar H+-ATPase (V-ATPase, sc-20943; Santa Cruz Biotechnology), and GFP (ab291–50; abcam, Cambridge, United Kingdom) were applied for 1 hour at room temperature followed by overnight incubation at +4°C. Primary antibodies were detected using fluorescent Cy2-, Cy3-, or Cy5-conjugated (Dianova, Hamburg, Germany) or HRP-conjugated secondary antibodies (Dako, Glostrup, Denmark). Double and triple staining was performed by sequential application of respective primary and secondary antibodies separated by washing steps. Fluorescent signals were evaluated by confocal microscopy using a Zeiss LSM 5 Exciter microscope with 40× and 63× objectives (N.A. 1.3/1.4) and processed with ZEN 2008 software. Light microscopy images were acquired with a LEICA DMRB microscope using a 100× objective (N.A. 1.30) and processed with the Axio Vision SE64 software. Three-dimensional structured illumination microscopy images were acquired using the OMX V4 Blaze system (GE Healthcare). Quantification of confocal signals by intensity was performed using Fiji software. Quantification of A-ICs, B-ICs, and PCs numbers in CNT/CD was performed in kidney sections concomitantly labeled for AQP2, pendrin, and V-ATPase or V1aR to identify PCs by their luminal AQP2 signal, B-ICs by their luminal pendrin and basolateral V-ATPase labeling, and A-ICs by their luminal V-ATPase staining.
Isolated Perfused Collecting Ducts
Adult (8–12 weeks) male C57BL/6J mice were euthanized by decapitation under isoflurane anesthesia followed by removal of the kidneys. CDs were dissected at the transition zone between cortex and outer medulla and processed for measurements of intracellular calcium concentrations ([Ca2+]i)or luminal pH. Four to six CDs were analyzed in each experimental setting. CDs were perfused with a double-barreled perfusion system of concentric pipettes in a temperate bath chamber. All measurements were performed in a pregassed (95% O2 and 5% CO2) bath solution. For [Ca2+]i measurements, CDs were incubated with 10 μmol/L Fura-2-AM in dissection solution for 1 hour at room temperature, and fluorescence intensities at 340 and 380 nm were monitored using an inverted microscope. After obtaining baseline values, CDs were treated with the V1aR agonist (A0–4-67; 50 or 100 nM) for 3 minutes followed by a washout period of 7–8 minutes and application of 50 nM AVP for 3 minutes; 340-to-380-nm signal ratios were calculated as an indicator of [Ca2+]i, and their peaks were compared between the treatments. For assessment of the luminal pH, CDs were perfused with 100 μmol/L 2ʹ,7ʹ-bis(carboxyethyl)-5(6ʹ)-carboxyfluorescein in luminal solution, and intensities of luminal fluorescence at 486 and 440 nm were monitored to calculate 486-to-440-nm signal ratio as a pH indicator. After equilibration for 5–10 minutes, CDs were treated with the V1aR agonist (100 nM for 4 minutes in the bath solution) followed by a washout for 10 minutes and application of 50 nM AVP for 4 minutes. Effects of the V1aR agonist and AVP on luminal pH were compared after normalization to the mean baseline values obtained during 30 seconds before the respective treatments.
Data are presented as means and SEM. We assumed normal distribution of our results on the basis of the experimental design. Results of animal experiments were evaluated by unpaired two-tailed t test to determine statistical significance. Ex vivo settings were analyzed by unpaired or paired t test and Kruskal–Wallis test followed by Dunn multiple comparisons test; P<0.05 was accepted as a statistically significant difference.
Segmental and Cellular V1aR Distribution in Rodent and Human Kidneys
To study the renal distribution of V1aR, we generated a polyclonal antibody to this receptor subtype by immunizing rabbits with a synthetic peptide corresponding to a C-terminal mouse V1aR portion (NH2-CKDSPKSSKSIRFIPVST-CONH2) with no significant homology to mouse V1bR or V2R but substantial homology to rat and human V1aR. Immunoperoxidase staining of mouse kidney produced basolateral or perinuclear to apical V1aR signal patterns in ICs of CNT and CD segments (Figure 1, A and B). Triple immunofluorescence labeling of V1aR, AQP2 as the luminal marker for PCs, and pendrin as the luminal marker for B-ICs allowed us to assign the basolateral V1aR signal to A-ICs and the perinuclear/subapical V1aR signal to B-ICs displaying apical pendrin (Figure 1, C–F; Supplemental Figure 1). Apical pendrin staining did not coincide with the V1aR signal, suggesting that the receptor did not reach the luminal membrane in B-ICs (Figure 1F). The distinct V1aR distribution patterns in A-ICs versus B-ICs were further verified using high-resolution three-dimensional structured illumination microscopy imaging (Figure 1, G and H; Supplemental Movie). In a small proportion of pendrin-positive cells, the V1aR signal was absent or showed either basolateral or diffuse localization (Figure 2, A–C). Similar to mice, rat and human kidneys showed basolateral or intracellular to subapical V1aR signals in AQP2-negative ICs (Figure 2, D–G). These localization data suggest basolateral V1aR-mediated effects of AVP in A-ICs, whereas B-ICs may be less responsive to AVP considering the virtual absence of V1aR in their plasma membrane. Other sites displaying renal V1aR immunoreactivity were MD cells showing a clear basolateral signal in mouse but not in rat or human kidney samples (Supplemental Figure 2). Renal arterioles and vasa recta showed a V1aR immunoreactive signal in the endothelium (Supplemental Figure 3). To further corroborate the localization data, we evaluated V1aR mRNA expression in microdissected mouse nephron segments using RT-PCR, which produced specific signals in CNT/CD and glomeruli with attached MD but not in proximal tubules, thick ascending limb, or distal convoluted tubule (Figure 1I, Supplemental Figure 4).
Specificity of the anti-V1aR antibody was confirmed by labeling of V1aR-deficient kidneys, which produced no significant signal in ICs or MD cells (Figure 3, A–D; Supplemental Figure 2, C and D). Immunoblotting with this antibody showed abundant signal of the expected molecular weight range in mouse liver samples, whereas brain and kidney signals were weak, possibly reflecting distinct numbers of V1aR-expressing cells in these organs (Figure 3E). Additional specificity tests used transfection experiments in cultured HEK293 cells lacking endogenous V1aR (Figure 3F). Transfection of mouse V1aR, FLAG-V1bR, or empty pcDNA3.1 plasmid followed by precipitation with protein G Sepharose gel and immunoblotting using anti-V1aR or anti-FLAG antibodies confirmed specificity of our antibody to V1aR and lack of crossreactivity with V1bR (Figure 3G). Along the same line, immunofluorescence labeling of HEK293 cells transfected with V1aR-GFP or control GFP-containing plasmid produced a clear plasma membrane–associated V1aR signal in the V1aR-GFP–transfected cells but not in controls (Figure 3, H–K).
Effects of V1aR Deletion on Distribution of CNT/CD Cell Types in Mouse Kidney
To test whether V1aR signaling affects the numerical proportions of IC types in CNT and CD, we quantified the numbers of PCs, A-ICs, and B-ICs in cortex and medulla of wild-type versus V1aR-deficient kidneys using triple labeling for AQP2, V-ATPase, and pendrin. The two genotypes showed similar percentages of PCs, A-ICs, and B-ICs, suggesting that V1aR is not essential for their proportional distribution in CNT and CD (Table 1).
Table 1. -
Distribution of collecting duct cell types in wild-type versus vasopressin V1a receptor–deficient kidneys.
Triple labeling for aquaporin 2 (AQP2), V-ATPase, and pendrin was performed to differentiate between the AQP2-positive principal cells (PCs), AQP2- and pendrin-negative type A intercalated cells (A-ICs), and pendrin-positive type B intercalated cells (B-ICs) in wild-type (WT; n=4) versus vasopressin V1a receptor–deficient (V1aR-KO) kidneys (n=4). At least ten to 20 tubules were quantified in each kidney zone; data are means.
Functional Effect of V1aR Stimulation versus Inhibition In Vivo
To mimic the physiologic effect of AVP binding to V1aR, we treated anesthetized and urinary bladder–catheterized mice with the V1aR agonist, AO-4–67 (2 μg/kg body weight), or vehicle. Compared with baseline values, urinary pH was significantly decreased after 20 minutes of the agonist administration (pH from 7.18 to 6.78; P<0.05) (Figure 4A) but unaffected in controls, indicating that selective stimulation of V1aR increases urinary acid load in vivo. To corroborate these results, we took advantage of AVP-deficient Brattleboro rats with central DI. Analysis of V1aR distribution in DI rats produced a signal pattern similar to that of normal rats (i.e., basolateral receptor presence in A-ICs versus diffused intracellular distribution in B-ICs) (Supplemental Figure 5, A–D). DI rats were treated with the V1aR agonist (200 ng/kg, 2 μg/kg, or 10 μg/kg body weight for 4 hours) to study changes of NAE in metabolic cages. The highest dose reduced urinary pH (from 7.40 to 6.71, 6.71, 6.56, and 6.53 after 1, 2, 3, and 4 hours, respectively; P<0.01) and HCO3− levels (−82%, −92%, and −85% after 2, 3, and 4 hours, respectively; P<0.001). NH4+ excretion was not significantly altered, but NAE showed significant increases (+176%, +248%, and +98% after 2, 3, and 4 hours, respectively) (Figure 4, C–F). The lower agonist doses produced only minor changes in the urine but led to a significant increase in plasma HCO3− without concomitant change of pH, likely reflecting the spared bicarbonate (27.6 versus 29.2 mmol/L; P<0.01) (Supplemental Figure 5E).
To study the role of V1aR during increased acid load, we induced metabolic acidosis in mice by administering 0.28 M NH4Cl with chow for 3 days.33 The resulting metabolic acidosis was confirmed by changes in plasma pH, HCO3−, and Cl− levels (Supplemental Figure 6). Intracellular V1aR distribution and proportions of V1aR-expressing ICs were unaffected in acidotic mice (Supplemental Figure 7, Supplemental Tables 1–4). Acute application of a V1aR antagonist in acidotic, bladder-catheterized mice (CL-14–102; 2 mg/kg body weight intraperitoneally) produced no significant changes in urinary pH compared with vehicle treatment (Figure 4B).
Effects of V1aR Stimulation on V-ATPase in Primary Cell Culture
To address V1aR-mediated effects on urinary acidification, IMCD cells isolated from rat kidney papilla were studied.35 To this end, V-ATPase expression in V1aR-immunoreactive A-ICs was first identified in situ in their papillary environment using double labeling for AQP2 and V1aR or for AQP2 and V-ATPase; A-ICs revealed substantial luminal V-ATPase and basolateral V1aR signals as opposed to AQP2-positive PCs (Figure 5, A and B). B-ICs were typically absent from this kidney zone. Isolated IMCD cells grown to confluence on permeable support similarly expressed V1aR and V-ATPase in single A-ICs scattered among AQP2-positive PCs (Figure 5, C and D). V1aR staining was detected along the lateral cell borders, whereas V-ATPase staining was distributed over the entire cell. To evaluate the effects of V1aR stimulation on V-ATPase in A-ICs, IMCD cells were treated with the V1aR agonist (1 μM, 4 hours) or vehicle applied from the basolateral side. V-ATPase immunoreactive signal intensity was measured using confocal z stacks of individual A-ICs over 7.5 μm of apicobasal extension. Apical V-ATPase signal was substantially enhanced in the supranuclear region of cells receiving the agonist as demonstrated by intensity plotting (+93%; P<0.001) (Figure 6, A–F). These data suggest vasopressin-inducible, V1aR-mediated activation of V-ATPase in A-ICs.
Effects of V1aR Stimulation in Isolated Perfused Collecting Duct Segments
Cellular V1aR signaling depends on intracellular Ca2+ release. We, therefore, used isolated perfused outer medullary CDs from mouse kidney to measure [Ca2+]i and luminal pH using Ca2+- (Fura-2 AM) and pH-sensitive [2ʹ,7ʹ-bis(carboxyethyl)-5(6ʹ)-carboxyfluorescein] fluorescent dyes.36,37 Basolateral treatment of CDs with the V1aR agonist, A0–4-67, induced dose-dependent, transient increases of [Ca2+]i. The consecutive application of AVP exerted stronger effects compared with a low V1aR agonist dose (1.20-fold for 50 nM A0–4-67 versus 1.50-fold for 50 nM AVP; P<0.05) but was comparable with a higher dose of the V1aR agonist (1.40-fold for 100 nM A0–4-67 versus 1.45-fold for consecutive 50 nM AVP) (Figure 7, A–E). We, therefore, selected the higher agonist dose to evaluate effects on luminal pH. Here, basolateral application of A0–4-67 or AVP caused significant luminal acidification to a similar extent (−4.55% for 100 nM A0–4-67 versus −4.15% for consecutive 50 nM AVP; P<0.05 for each treatment versus respective baseline values) (Figure 7, F and G). Together, our data indicate that V1aR stimulation induces luminal H+ secretion in CDs, thus corroborating and extending the in vivo data.
In this study, we focused on the role of ICs in AVP-V1aR–mediated control of acid-base homeostasis in the CD system. Expression of V1aR in CNTs and CDs has been reported, yet cell type–specific characterization of its distribution remained uncertain.16,19,20,38,39 Earlier localization and functional studies suggested predominantly apical V1aR distribution in ICs or PCs.19,39–43 Indeed, AVP is filtered into the urine, where its concentration substantially exceeds plasma levels.5,27 Luminal effects of AVP on [Ca2+]i or transepithelial resistance have been established in the rabbit CD, whereas robust functional data from rodent or human kidneys are not available.40,43 Other studies demonstrated basolateral effects of V1aR stimulation in rabbits and rats, although the responsive cell types were not defined.41,44
In this work, we used a new anti-V1aR antibody to clarify segment- and cell type–specific aspects of V1aR distribution in mouse, rat, and human kidneys. The clear basolateral, membrane-bound V1aR signal in A-IC is suggestive of their responsiveness to plasma AVP, whereas the intracellular signal without significant plasma membrane association in B-ICs points to their poor AVP sensitivity compared with A-ICs.
Our functional studies further support direct effects of AVP in A-ICs. Stimulation of the AVP-V1aR axis in vivo by a V1aR agonist decreased urinary pH and increased NAE in AVP-deficient DI rats, which is consistent with activation of A-ICs. Because of the lack of endogenous AVP, DI rats were assumed to exhibit particularly strong responses to AVP receptor agonists.45 However, a relatively high dose of V1aR agonist was required to induce significant effects in this model, which may be related to altered urinary buffering capacities in the state of extreme diuresis or volume depletion in Brattleboro rats. V1aR activation may further elicit indirect effects via potentiation of aldosterone action.19 Bladder catheterization in mice has permitted particularly robust recordings of the urinary acid-base status.37 To over-ride the effects of endogenous AVP, we applied a saturating dose of V1aR agonist in mice, which resulted in rapid decreases of urinary pH, likely due to activation of A-ICs.
These results were extended in our ex vivo models using isolated perfused CDs or cultured IMCD cells. Because V1aR signals via intracellular calcium release,46 we have detected [Ca2+]i parallel to evaluation of luminal pH in isolated CDs. Basolateral application of the agonist in isolated perfused CDs reduced luminal pH and elicited transient [Ca2+]i increases, which is in line with the assumed recruitment of intracellular calcium as a second messenger.47 Because virtually all epithelial cells of isolated tubules exhibited increased [Ca2+]i in response to the agonist, we suggest that direct effects on A-ICs may elicit paracrine signaling events affecting neighboring PCs.21 Administration of AVP produced similar increases of [Ca2+]i and luminal pH reduction as the V1aR agonist, suggesting that urinary acidification is not an artifact of selective V1aR stimulation but rather, a physiologic AVP function. These results also suggest that concomitant V2R activation in PCs does not elicit inhibiting paracrine effects on V1aR-mediated H+ secretion. The model of isolated perfused CDs thus permitted us to detect significant effects of basolateral V1aR activation on luminal pH in the absence of systemic factors or contribution of upstream nephron segments. As to mechanisms involved, our experiments in cultured IMCD cells suggest stimulation of V-ATPase via its luminal trafficking.48 Cultured IMCD cells have been previously established as a model of PCs function.35 Here, we made use of these cells to study regulation of A-ICs, which are present in IMCD along with PCs.21,49 Growing IMCD cells on permeable support enabled basolateral application of the V1aR agonist. The resulting signal increases of apical V-ATPase in A-ICs suggested activation of the proton pump via luminal trafficking, although we cannot exclude changes in biosynthesis rate or protein stability.48 Other H+-secreting proteins may be affected in parallel.50 Previous work showed that aldosterone also activates V-ATPase in A-ICs via its luminal trafficking in a Ca2+-dependent manner, which may reflect involvement of a common downstream mediator, such as protein kinase C.51
V1aR-deficient mice exhibit dRTA,19,52 which led us to test the role of V1aR in renal adaptation to acid load using an NH4Cl-rich diet.33 Previous work reported increased V1aR mRNA expression in ICs of acid-loaded mice.20 Our data showed unaltered cellular V1aR distribution upon acid loading, suggesting an absence of compensatory changes at this level. V1aR antagonist had no effects on urinary pH in this model, suggesting a minor role for V1aR signaling in adaptations to metabolic acidosis. Local pH-dependent modulation of V1aR signaling in ICs may be considered as well (i.e., H+ secretion may be stimulated by local alkaline pH but inhibited by acidic pH).53
The nondetectable membrane-associated V1aR signal in B-ICs, despite intracellular receptor accumulation, suggests that AVP does not elicit direct effects in this cell type. However, minor luminal surface expression of the receptor below the given antibody detection limit may still be functional considering the naturally high urinary concentrations of AVP.5,27 In this case, rapid binding of AVP to luminal V1aR followed by receptor internalization could potentially explain its intracellular accumulation in B-ICs.5,54,55 However, this was not confirmed in AVP-deficient DI rats, which also lacked a membrane-bound receptor signal.
In this context, neither V1aR deletion in mice nor V1aR knockdown in cultured ICs altered the expression of pendrin, the Cl−/HCO3− exchanger of B-ICs.19 In contrast, in V2R-deficient mice, pendrin was suppressed in these cells.56 Along the same line, the V2R agonist desmopressin increased renal pendrin expression in DI rats.23 Because V2R is not expressed in ICs,57 these effects may have been elicited by neighboring V2R-expressing PCs via paracrine interactions involving PG, purinergic signaling, or other pathways.21,26,58,59 Available data thus do not provide evidence for a direct sensitivity of B-ICs to AVP but suggest paracrine crosstalk between different CD cell types.40,42,60
Mechanisms underlying the distinct V1aR distribution patterns in A-ICs and B-ICs are not clear at present. A recent transcriptome analysis demonstrated similar abundances of V1aR mRNA expression in both cell types, suggesting that post-translational regulation determines the individual destination of the receptor.61 Extrinsic factors may potentially modulate functional properties of ICs, including AVP signaling mechanisms; adaptive conversion of B-ICs to A-ICs or proliferation of A-ICs serves to prevent dRTA.26,52,62 The role of A-ICs in the renal acid-base handling is illustrated by pronounced dRTA in hensin-deficient mice lacking this IC-type due to impaired hensin-dependent conversion of B-ICs to A-ICs.63 Although our data suggest functional V1aR-dependent activation of A-ICs, we obtained no evidence for a role of V1aR in IC acquisition of distinct phenotypes.
Apart from direct effects on A-ICs, AVP may affect renal acid-base handling via stimulation of RAAS.18,25,64 RAAS components exert permissive effects on functions in all CD cell types.21,58,65,66 Angiotensin II and aldosterone stimulate V-ATPase in A-ICs, thus increasing luminal H+ secretion, whereas aldosterone additionally induces pendrin in B-ICs, which facilitates the function of the epithelial sodium channel in PCs.65–69 The AVP-dependent modulation of RAAS may take place at the level of hypothalamus via stimulation of adrenocorticotropic hormone release as reported in rats.70 In mice, effects of AVP on RAAS may additionally be mediated by V1aR in MD cells, because V1aR-deficient mice exhibited hyporeninemia and low BP.18
In summary, this study extends information on V1aR distribution in rodent and human kidneys and provides several lines of evidence for AVP-induced, V1aR-mediated urinary H+ secretion by A-ICs.
We thank Kerstin Riskowsky and Frauke Grams for excellent technical assistance; Martin Thomson (Charité–Universitätsmedizin Berlin), Finn Antrobus and Yvonne Giesecke (both University of St. Andrews) for proofreading the manuscript; Prof. Carsten Wagner (Zurich, Switzerland) for providing us with the antipendrin antibody; Prof. Maurice Manning (Toledo, OH) for providing us with the V1aR agonist A0-4-67 and with the V1aR antagonist CL-14-102; and Yvonne Giesecke for help with cell quantification.
The work was supported by the Charité–Universitätsmedizin Berlin and Deutsche Forschungsgemeinschaft grants MU2924/2-2 and BA700/22-2. Mr. Torsten Giesecke received a joint research grant from the Berlin Institute of Health and the Charité–Universitätsmedizin Berlin. The work of Dr. Gimber was supported by the Advanced Medical BioImaging Core Facility of the Charite–Universitätsmedizin Berlin and Deutsche Forschungsgemeinschaft grant SFB958/Z02 (to Dr. Jan Schmoranzer). Dr. Paliege was supported by the Charité Clinical Scientist Program financed by the Charité–Universitätsmedizin Berlin and the Berlin Institute of Health.
This article contains the following supplemental material online at http://jasn.asnjournals.org/lookup/suppl/doi:10.1681/ASN.2018080816/-/DCSupplemental.
Supplemental Figure 1. Overview of V1aR distribution in mouse kidney.
Supplemental Figure 2. Distribution of V1aR in macula densa cells.
Supplemental Figure 3. V1aR distribution in renal vasculature of mouse kidney.
Supplemental Figure 4. Evaluation of microdissected nephron segments.
Supplemental Figure 5. V1aR distribution in Brattleboro rats and effects of the V1aR agonist.
Supplemental Figure 6. Verification of metabolic acidosis in mice.
Supplemental Figure 7. Effects of metabolic acidosis on cellular V1aR distribution.
Supplemental Material. Complete methods: extended methods description.
Supplemental Movie. 3D reconstruction of structured illumination microscopy (3D-SIM) showing subcellular distribution of V1aR in mouse kidney.
Supplemental Table 1. Proportion of collecting duct cell types in control versus acidotic mice.
Supplemental Table 2. Evaluation of net acid excretion in human urine.
Supplemental Table 3. Evaluation of net acid excretion in Brattleboro rat urine.
Supplemental Table 4. Measurement of the standard for experiments in Supplemental Tables 2 and 3.
1. Kortenoeven ML, Pedersen NB, Rosenbaek LL, Fenton RA: Vasopressin regulation of sodium transport in the distal nephron and collecting duct. Am J Physiol Renal Physiol 309: F280–F299, 201526041443
2. Gunaratne R, Braucht DW, Rinschen MM, Chou CL, Hoffert JD, Pisitkun T, et al.: Quantitative phosphoproteomic analysis reveals cAMP/vasopressin-dependent signaling pathways in native renal thick ascending limb cells. Proc Natl Acad Sci U S A 107: 15653–15658, 201020713729
3. Douglass J, Gunaratne R, Bradford D, Saeed F, Hoffert JD, Steinbach PJ, et al.: Identifying protein kinase target preferences using mass spectrometry. Am J Physiol Cell Physiol 303: C715–C727, 201222723110
4. Robben JH, Knoers NV, Deen PM: Cell biological aspects of the vasopressin type-2 receptor and aquaporin 2 water channel in nephrogenic diabetes insipidus. Am J Physiol Renal Physiol 291: F257–F270, 200616825342
5. Bankir L: Antidiuretic action of vasopressin: Quantitative aspects and interaction between V1a and V2 receptor-mediated effects. Cardiovasc Res 51: 372–390, 200111476728
6. Bankir L, Fernandes S, Bardoux P, Bouby N, Bichet DG: Vasopressin-V2 receptor stimulation reduces sodium excretion in healthy humans. J Am Soc Nephrol 16: 1920–1928, 200515888562
7. Fernandes S, Bruneval P, Hagege A, Heudes D, Ghostine S, Bouby N: Chronic V2 vasopressin receptor stimulation increases basal blood pressure and exacerbates deoxycorticosterone acetate-salt hypertension. Endocrinology 143: 2759–2766, 200212072411
8. Luft FC: Vasopressin, urine concentration, and hypertension: A new perspective on an old story. Clin J Am Soc Nephrol 2: 196–197, 200717699405
9. Bardoux P, Bruneval P, Heudes D, Bouby N, Bankir L: Diabetes-induced albuminuria: Role of antidiuretic hormone as revealed by chronic V2 receptor antagonism in rats. Nephrol Dial Transplant 18: 1755–1763, 200312937221
10. Bardoux P, Martin H, Ahloulay M, Schmitt F, Bouby N, Trinh-Trang-Tan MM, et al.: Vasopressin contributes to hyperfiltration, albuminuria, and renal hypertrophy in diabetes mellitus: Study in vasopressin-deficient Brattleboro rats. Proc Natl Acad Sci U S A 96: 10397–10402, 199910468619
11. Rinschen MM, Schermer B, Benzing T: Vasopressin-2 receptor signaling and autosomal dominant polycystic kidney disease: From bench to bedside and back again. J Am Soc Nephrol 25: 1140–1147, 201424556353
12. Bankir L, Bouby N, Ritz E: Vasopressin: A novel target for the prevention and retardation of kidney disease? Nat Rev Nephrol 9: 223–239, 201323438973
13. Okada H, Suzuki H, Kanno Y, Yamamura Y, Saruta T: Chronic and selective vasopressin blockade in spontaneously hypertensive rats. Am J Physiol 267: R1467–R1471, 19947810754
14. Okada H, Suzuki H, Kanno Y, Saruta T: Effects of novel, nonpeptide vasopressin antagonists on progressive nephrosclerosis in rats. J Cardiovasc Pharmacol 25: 847–852, 19957630164
15. Perico N, Zoja C, Corna D, Rottoli D, Gaspari F, Haskell L, et al.: V1/V2 Vasopressin receptor antagonism potentiates the renoprotection of renin-angiotensin system inhibition in rats with renal mass reduction. Kidney Int 76: 960–967, 200919625993
16. Ostrowski NL, Young WS 3rd, Knepper MA, Lolait SJ: Expression of vasopressin V1a and V2 receptor messenger ribonucleic acid in the liver and kidney of embryonic, developing, and adult rats. Endocrinology 133: 1849–1859, 19938404628
17. Serradeil-Le Gal C, Raufaste D, Marty E, Garcia C, Maffrand JP, Le Fur G: Autoradiographic localization of vasopressin V1a receptors in the rat kidney using [3
H]-SR 49059. Kidney Int 50: 499–505, 19968840278
18. Aoyagi T, Izumi Y, Hiroyama M, Matsuzaki T, Yasuoka Y, Sanbe A, et al.: Vasopressin regulates the renin-angiotensin-aldosterone system via V1a receptors in macula densa cells. Am J Physiol Renal Physiol 295: F100–F107, 200818448596
19. Izumi Y, Hori K, Nakayama Y, Kimura M, Hasuike Y, Nanami M, et al.: Aldosterone requires vasopressin V1a receptors on intercalated cells to mediate acid-base homeostasis. J Am Soc Nephrol 22: 673–680, 201121415155
20. Yasuoka Y, Kobayashi M, Sato Y, Zhou M, Abe H, Okamoto H, et al.: The intercalated cells of the mouse kidney OMCD(is) are the target of the vasopressin V1a receptor axis for urinary acidification. Clin Exp Nephrol 17: 783–792, 201323456233
21. Roy A, Al-bataineh MM, Pastor-Soler NM: Collecting duct intercalated cell function and regulation. Clin J Am Soc Nephrol 10: 305–324, 201525632105
22. Bichara M, Mercier O, Houillier P, Paillard M, Leviel F: Effects of antidiuretic hormone on urinary acidification and on tubular handling of bicarbonate in the rat. J Clin Invest 80: 621–630, 19873624481
23. Amlal H, Sheriff S, Faroqui S, Ma L, Barone S, Petrovic S, et al.: Regulation of acid-base transporters by vasopressin in the kidney collecting duct of Brattleboro rat. Am J Nephrol 26: 194–205, 200616699257
24. Petrovic S, Amlal H, Sun X, Karet F, Barone S, Soleimani M: Vasopressin induces expression of the Cl-/HCO3- exchanger SLC26A7 in kidney medullary collecting ducts of Brattleboro rats. Am J Physiol Renal Physiol 290: F1194–F1201, 200616352747
25. Möhring J, Kohrs G, Möhring B, Petri M, Homsy E, Haack D: Effects of prolonged vasopressin treatment in Brattleboro rats with diabetes insipidus. Am J Physiol 234: F106–F111, 1978623299
26. Schwartz GJ, Gao X, Tsuruoka S, Purkerson JM, Peng H, D’Agati V, et al.: SDF1 induction by acidosis from principal cells regulates intercalated cell subtype distribution. J Clin Invest 125: 4365–4374, 201526517693
27. El-Farhan N, Hampton D, Penney M: Measurement of arginine vasopressin. Methods Mol Biol 1065: 129–139, 201323996361
28. Manning M, Sawyer WH: Design and uses of selective agonistic and antagonistic analogs of the neuropeptides oxytocin and vasopressin. Trends Neurosci 7: 6–9, 1984
29. Szczepanska-Sadowska E, Stepniakowski K, Skelton MM, Cowley AW Jr.: Prolonged stimulation of intrarenal V1 vasopressin receptors results in sustained hypertension. Am J Physiol 267: R1217–R1225, 19947977848
30. Chan JC: The rapid determination of urinary titratable acid and ammonium and evaluation of freezing as a method of preservation. Clin Biochem 5: 94–98, 19725039597
31. Chan WY, Wo NC, Cheng LL, Manning M: Isosteric substitution of Asn5 in antagonists of oxytocin and vasopressin leads to highly selective and potent oxytocin and V1a receptor antagonists: New approaches for the design of potential tocolytics for preterm labor. J Pharmacol Exp Ther 277: 999–1003, 19968627583
32. Shirley DG, Walter MF, Keeler BD, Waters NJ, Walter SJ: Selective blockade of oxytocin and vasopressin V(1a) receptors in anaesthetised rats: Evidence that activation of oxytocin receptors rather than V(1a) receptors increases sodium excretion. Nephron Physiol 117: 21–26, 201121071981
33. Nowik M, Kampik NB, Mihailova M, Eladari D, Wagner CA: Induction of metabolic acidosis with ammonium chloride (NH4Cl) in mice and rats--species differences and technical considerations. Cell Physiol Biochem 26: 1059–1072, 201021220937
34. Kashiwazaki A, Fujiwara Y, Tsuchiya H, Sakai N, Shibata K, Koshimizu TA: Subcellular localization and internalization of the vasopressin V1B receptor. Eur J Pharmacol 765: 291–299, 201526318147
35. Faust D, Geelhaar A, Eisermann B, Eichhorst J, Wiesner B, Rosenthal W, et al.: Culturing primary rat inner medullary collecting duct cells [published online ahead of print June 21, 2013]. J. Vis. Exp 10.3791/50366
36. Ankorina-Stark I, Haxelmans S, Schlatter E: Functional evidence for the regulation of cytosolic Ca2+ activity via V1A-receptors and beta-adrenoceptors in rat CCD. Cell Calcium 21: 163–171, 19979132299
37. de Bruijn PI, Larsen CK, Frische S, Himmerkus N, Praetorius HA, Bleich M, et al.: Furosemide-induced urinary acidification is caused by pronounced H+ secretion in the thick ascending limb. Am J Physiol Renal Physiol 309: F146–F153, 201525995110
38. Carmosino M, Brooks HL, Cai Q, Davis LS, Opalenik S, Hao C, et al.: Axial heterogeneity of vasopressin-receptor subtypes along the human and mouse collecting duct. Am J Physiol Renal Physiol 292: F351–F360, 200716835408
39. Gonzalez CB, Figueroa CD, Reyes CE, Caorsi CE, Troncoso S, Menzel D: Immunolocalization of V1 vasopressin receptors in the rat kidney using anti-receptor antibodies. Kidney Int 52: 1206–1215, 19979350643
40. Ando Y, Tabei K, Asano Y: Luminal vasopressin modulates transport in the rabbit cortical collecting duct. J Clin Invest 88: 952–959, 19911885780
41. Naruse M, Yoshitomi K, Hanaoka K, Imai M, Kurokawa K: Electrophysiological study of luminal and basolateral vasopressin in rabbit cortical collecting duct. Am J Physiol 268: F20–F29, 19957840244
42. Amorim JB, Malnic G: V(1) receptors in luminal action of vasopressin on distal K(+) secretion. Am J Physiol Renal Physiol 278: F809–F816, 200010807593
43. Ikeda M, Yoshitomi K, Imai M, Kurokawa K: Cell Ca2+ response to luminal vasopressin in cortical collecting tubule principal cells. Kidney Int 45: 811–816, 19948196283
44. Rivarola V, Ford P, del Pilar Flamenco M, Galizia L, Capurro C: Arginine-vasopressin modulates intracellular pH via V1 and V2 receptors in renal collecting duct cells. Cell Physiol Biochem 20: 549–558, 200717762181
45. Mutig K, Paliege A, Kahl T, Jöns T, Müller-Esterl W, Bachmann S: Vasopressin V2 receptor expression along rat, mouse, and human renal epithelia with focus on TAL. Am J Physiol Renal Physiol 293: F1166–F1177, 200717626156
46. Briley EM, Lolait SJ, Axelrod J, Felder CC: The cloned vasopressin V1a receptor stimulates phospholipase A2, phospholipase C, and phospholipase D through activation of receptor-operated calcium channels. Neuropeptides 27: 63–74, 19947969820
47. Tordjmann T, Berthon B, Jacquemin E, Clair C, Stelly N, Guillon G, et al.: Receptor-oriented intercellular calcium waves evoked by vasopressin in rat hepatocytes. EMBO J 17: 4695–4703, 19989707428
48. Breton S, Brown D: New insights into the regulation of V-ATPase
-dependent proton secretion. Am J Physiol Renal Physiol 292: F1–F10, 200717032935
49. Madsen KM, Clapp WL, Verlander JW: Structure and function of the inner medullary collecting duct. Kidney Int 34: 441–454, 19883059025
50. Gumz ML, Lynch IJ, Greenlee MM, Cain BD, Wingo CS: The renal H+-K+-ATPases: Physiology, regulation, and structure. Am J Physiol Renal Physiol 298: F12–F21, 201019640897
51. Winter C, Kampik NB, Vedovelli L, Rothenberger F, Paunescu TG, Stehberger PA, et al.: Aldosterone stimulates vacuolar H(+)-ATPase activity in renal acid-secretory intercalated cells mainly via a protein kinase C-dependent pathway. Am J Physiol Cell Physiol 301: C1251–C1261, 201121832245
52. Welsh-Bacic D, Nowik M, Kaissling B, Wagner CA: Proliferation of acid-secretory cells in the kidney during adaptive remodelling of the collecting duct. PLoS One 6: e25240, 201122039408
53. Brown D, Wagner CA: Molecular mechanisms of acid-base sensing by the kidney. J Am Soc Nephrol 23: 774–780, 201222362904
54. Bowen-Pidgeon D, Innamorati G, Sadeghi HM, Birnbaumer M: Arrestin effects on internalization of vasopressin receptors. Mol Pharmacol 59: 1395–1401, 200111353798
55. Innamorati G, Le Gouill C, Balamotis M, Birnbaumer M: The long and the short cycle. Alternative intracellular routes for trafficking of G-protein-coupled receptors. J Biol Chem 276: 13096–13103, 200111150299
56. Schliebe N, Strotmann R, Busse K, Mitschke D, Biebermann H, Schomburg L, et al.: V2 vasopressin receptor deficiency causes changes in expression and function of renal and hypothalamic components involved in electrolyte and water homeostasis. Am J Physiol Renal Physiol 295: F1177–F1190, 200818715941
57. Mutig K, Borowski T, Boldt C, Borschewski A, Paliege A, Popova E, et al.: Demonstration of the functional impact of vasopressin signaling in the thick ascending limb by a targeted transgenic rat approach. Am J Physiol Renal Physiol 311: F411–F423, 201627306979
58. Pearce D, Soundararajan R, Trimpert C, Kashlan OB, Deen PM, Kohan DE: Collecting duct principal cell transport processes and their regulation. Clin J Am Soc Nephrol 10: 135–146, 201524875192
59. Gueutin V, Vallet M, Jayat M, Peti-Peterdi J, Cornière N, Leviel F, et al.: Renal β-intercalated cells maintain body fluid and electrolyte balance. J Clin Invest 123: 4219–4231, 201324051376
60. Barreto-Chaves ML, de Mello-Aires M: Luminal arginine vasopressin stimulates Na(+)-H+ exchange and H(+)-ATPase in cortical distal tubule via V1 receptor. Kidney Int 52: 1035–1041, 19979328942
61. Chen L, Lee JW, Chou CL, Nair AV, Battistone MA, Păunescu TG, et al.: Transcriptomes of major renal collecting duct cell types in mouse identified by single-cell RNA-seq. Proc Natl Acad Sci U S A 114: E9989–E9998, 201729089413
62. Purkerson JM, Tsuruoka S, Suter DZ, Nakamori A, Schwartz GJ: Adaptation to metabolic acidosis and its recovery are associated with changes in anion exchanger distribution and expression in the cortical collecting duct. Kidney Int 78: 993–1005, 201020592712
63. Gao X, Eladari D, Leviel F, Tew BY, Miró-Julià C, Cheema FH, et al.: Deletion of hensin/DMBT1 blocks conversion of beta- to alpha-intercalated cells and induces distal renal tubular acidosis. Proc Natl Acad Sci U S A 107: 21872–21877, 201021098262
64. Kinter LB, Shier D, Flamenbaum W, Beeuwkes R 3rd: The renin angiotensin system in conscious Brattleboro strain rats. Ren Physiol 5: 278–285, 19827178644
65. Pech V, Zheng W, Pham TD, Verlander JW, Wall SM: Angiotensin II activates H+-ATPase in type A intercalated cells. J Am Soc Nephrol 19: 84–91, 200818178800
66. Mohebbi N, Perna A, van der Wijst J, Becker HM, Capasso G, Wagner CA: Regulation of two renal chloride transporters, AE1 and pendrin, by electrolytes and aldosterone. PLoS One 8: e55286, 201323383138
67. Wall SM: Renal intercalated cells and blood pressure regulation. Kidney Res Clin Pract 36: 305–317, 201729285423
68. Verlander JW, Hassell KA, Royaux IE, Glapion DM, Wang ME, Everett LA, et al.: Deoxycorticosterone upregulates PDS (Slc26a4) in mouse kidney: Role of pendrin in mineralocorticoid-induced hypertension. Hypertension 42: 356–362, 200312925556
69. Pech V, Wall SM, Nanami M, Bao HF, Kim YH, Lazo-Fernandez Y, et al.: Pendrin gene ablation alters ENaC subcellular distribution and open probability. Am J Physiol Renal Physiol 309: F154–F163, 201525972513
70. Brudieux R, Krifi MN, Laulin JP: Release of aldosterone and corticosterone from the adrenal cortex of the Brattleboro rat in response to administration of ACTH. J Endocrinol 111: 375–381, 19863027224